Download Vasopressin-stimulated Ca2 spiking in vascular smooth muscle cells

Am J Physiol Heart Circ Physiol
280: H2658–H2664, 2001.
Vasopressin-stimulated Ca2⫹ spiking in
vascular smooth muscle cells involves phospholipase D
YANXIA LI,1 AARON J. SHIELS,2 GARY MASZAK,2 AND KENNETH L. BYRON2
Department of Physiology, Loyola University, Chicago, 60626; and 2Department of Medicine,
Stritch School of Medicine, Maywood, Illinois 60153
1
Received 22 February 2000; accepted in final form 10 January 2001
A7r5; lipid hydrolysis; protein kinase C; signal transduction
of contraction of
vascular smooth muscle (VSM). Vasoconstrictor hormones such as [Arg8]vasopressin (AVP), which bind to
heptahelical G protein-coupled receptors, are thought
to exert vasoconstrictor actions via activation of phospholipase C (PLC). The product of this pathway, inositol 1,4,5-trisphosphate [Ins(1,4,5)P3], increases the cytosolic free Ca2⫹ concentration ([Ca2⫹]i) via the release
of Ca2⫹ from intracellular Ca2⫹ stores. The concentration of AVP needed to half-maximally increase
Ins(1,4,5)P3 and release intracellular Ca2⫹ (⬃2 nM) is
much higher than the vasoconstrictor concentrations of
AVP normally found in the systemic circulation (10–
100 pM). This raises a question as to whether the
CALCIUM ION IS THE PRIMARY MEDIATOR
Address for reprint requests and other correspondence: K. L.
Byron, Cardiovascular Institute, Loyola Univ. Medical Center, 2160
South First Ave., Maywood, IL 60153 (E-mail: [email protected]).
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release of intracellular Ca2⫹ stores can account for the
vasoconstrictor effects of AVP. We have recently shown
(2) that physiological concentrations of AVP (10–500
pM) stimulate oscillations of [Ca2⫹]i (Ca2⫹ spikes) that
increase in frequency with increasing AVP concentration ([AVP]) in A7r5 vascular smooth muscle cells.
These Ca2⫹ spikes arise due to Ca2⫹-dependent action
potentials. Hence, in contrast to InsP3-mediated release of intracellular Ca2⫹, the Ca2⫹ spikes have a
strict requirement for extracellular Ca2⫹ and are abolished by blockers of voltage-sensitive Ca2⫹ channels.
AVP-stimulated Ca2⫹ spiking in A7r5 cells was previously found (2) to be blocked by a putative inhibitor of
phospholipase A2 (PLA2), ONO-RS-082, leading to the
conclusion that PLA2 may mediate this effect of AVP.
The present study examines in more detail the lipid
products of PLA2 and the action of ONO-RS-082. The
results suggest that phospholipase D (PLD) rather
than PLA2 may mediate the stimulation of Ca2⫹ spiking by AVP.
MATERIALS AND METHODS
Cell culture. A7r5 cells were cultured as described previously (3). Cells were subcultured onto rectangular 9 ⫻ 22-mm
(no. 11⁄2) glass coverslips or plastic tissue-culture dishes
(Corning). Confluent cells (passages 10–30) were used 2–5
days after plating. The health and phenotype of the cells
were verified routinely by examining [Ca2⫹]i responses to 100
pM AVP. Similar responses were obtained with all cell passages tested including those used for all of the biochemical
assays.
Loading cells with fura 2. Coverslips were washed twice
with control medium (in mM: 135 NaCl, 5.9 KCl, 1.5 CaCl2,
1.2 MgCl2, 11.5 glucose, and 11.6 HEPES/NaOH; pH 7.3) and
then incubated in the same medium with 2 ␮M fura 2-acetoxymethyl ester (AM), 0.1% bovine serum albumin (BSA),
and 0.025% Pluronic F-127 detergent (20) for 90–120 min at
room temperature (20–23°C). After loading, the cells were
washed twice and incubated in control medium for 1–5 h
before the start of the experiment. This final incubation
allowed for complete hydrolysis of fura 2-AM as assessed by
the shift in the fluorescence spectrum (24). As noted previously (3), this dye-loading protocol appeared not to adversely
affect the cells. About 95% of the fura 2 was released from the
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0363-6135/01 $5.00 Copyright © 2001 the American Physiological Society
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Li, Yanxia, Aaron J. Shiels, Gary Maszak, and Kenneth L. Byron. Vasopressin-stimulated Ca2⫹ spiking in
vascular smooth muscle cells involves phospholipase D.
Am J Physiol Heart Circ Physiol 280: H2658–H2664, 2001.—
Physiological concentrations of [Arg8]vasopressin (AVP; 10–
500 pM) stimulate oscillations of cytosolic free Ca2⫹ concentration (Ca2⫹ spikes) in A7r5 vascular smooth muscle cells.
We previously reported that this effect of AVP was blocked by
a putative phospholipase A2 (PLA2) inhibitor, ONO-RS-082
(5 ␮M). In the present study, the products of PLA2, arachidonic acid (AA), and lysophospholipids were found to be
ineffective in stimulating Ca2⫹ spiking, and inhibitors of AA
metabolism did not prevent AVP-stimulated Ca2⫹ spiking.
Thin layer chromatography was used to monitor the release
of AA and phosphatidic acid (PA), which are the products of
PLA2 and phospholipase D (PLD), respectively. AVP (100
pM) stimulated both AA and PA formation, but only PA
formation was inhibited by ONO-RS-082 (5 ␮M). Exogenous
PLD (type VII; 2.5 U/ml) stimulated Ca2⫹ spiking equivalent
to the effect of 100 pM AVP. AVP stimulated transphosphatidylation of 1-butanol (a PLD-catalyzed reaction) but not
2-butanol, and 1-butanol (but not 2-butanol) completely prevented AVP-stimulated Ca2⫹ spiking. Protein kinase C
(PKC) inhibition, which completely prevents AVP-stimulated
Ca2⫹ spiking, did not inhibit AVP-stimulated phosphatidylbutanol formation. These results suggest that AVP-stimulated Ca2⫹ spiking depends on activation of PLD rather than
PLA2 and that PKC activation may be downstream of PLD in
the signaling cascade.
ROLE OF PLD IN AVP-STIMULATED CA2⫹ SPIKING IN A7r5 CELLS
four experiments were compared by one-way ANOVA and
were considered statistically significant at P ⬍ 0.05.
PLD activation assayed by TLC. A7r5 cells were grown to
confluence in 100-mm petri dishes and labeled for 24 h with
[3H]palmitic acid (4 ␮Ci in 6 ml of DMEM supplemented with
3.5% fetal bovine serum at 37°C). The cells were pretreated
for 3 h in control medium supplemented with 0.1% fatty
acid-free BSA, and 0.2% 1-butanol was added during the last
4 min of the pretreatment. The cells were then treated with
AVP ⫹ 0.2% 1-butanol in 10 ml of medium for 20 min at 37°C.
At the end of this incubation, the medium was removed and
1.5 ml of ice-cold methanol was added to the dishes. After 20
min at 4°C, the cells were scraped from the dish and extracted according to the methods of Bligh and Dyer (1). The
extracted lipids were dried with N2 gas, resuspended in
chloroform with unlabeled PA and dipalmitoyl phosphatidylbutanol, and spotted on Whatman 60A-LKD TLC plates. The
plates were developed sequentially in three solvent mixtures
(18) to assure separation of phosphatidylbutanol from PA,
phosphatidylcholine, and neutral lipids. The locations of the
lipids on the plate were detected using molybdenum blue
reagent, and bands corresponding to locations of the unlabeled phosphatidylbutanol standard were scraped into scintillation vials and counted in an LKB-1209 liquid scintillation counter. Results were compared by one-way ANOVA and
were considered statistically significant at P ⬍ 0.05.
Materials. Cell-culture media were from GIBCO-BRL or
MediaTech. Fura 2-AM, fura 2 pentapotassium salt, and
Pluronic F-127 were from Molecular Probes; EGTA (puriss.
grade) was from Fluka Chemical; [3H]AA and [3H]palmitic
acid were from American Radiolabeled Chemicals; and dipalmitoylphosphatidyl butanol was from Avanti Polar Lipids. ONO-RS-082 was from Biomol. AA from several sources
was tested (Sigma, Calbiochem, Fluka, and Biomol) and was
used fresh or after storage under N2 gas. All other chemicals
including AVP and PLD type VII were from Sigma.
RESULTS
AA metabolism. We previously found that the putative PLA2 inhibitor ONO-RS-082 blocked both AVPstimulated Ca2⫹ spiking and AVP-stimulated release
of [3H]AA from A7r5 cells (2). Our interpretation of
these findings was that PLA2 might be involved in
AVP-stimulated Ca2⫹ spiking. To determine whether
the product of PLA2, AA, is important for stimulation
of Ca2⫹ spiking, fura 2-loaded A7r5 cell monolayers
were treated with varying concentrations of AA (see
Fig. 1). AA added to the medium at concentrations
from 1 nM to 50 ␮M did not stimulate Ca2⫹ spiking. AA
from several sources was tested and consistently failed
to induce spiking, although at concentrations ⱖ20 ␮M
a gradual increase in baseline [Ca2⫹]i was observed
(see Fig. 1A).
It is possible that AA cleaved from membrane phospholipids exerts a local effect which is not achieved by
addition of exogenous AA to the extracellular medium.
AA produced via PLA2 may be metabolized by cyclooxygenase, lipoxygenase, and cytochrome P-450 pathways to produce a variety of other signaling molecules.
A number of pharmacological inhibitors of these AA
metabolic pathways were tested for effects on AVPstimulated Ca2⫹ spiking. Inhibitors of cyclooxygenase
(10 ␮M indomethacin or 20–50 ␮M ibuprofen), lipoxygenase (10 ␮M 5,8,11-eicosatriynoic acid, 1–10 ␮M
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cells within 3 min of saponin (50 ␮g/ml) addition which
suggests that ⬃95% of the dye was in the cytosol.
[Ca2⫹]i measurements. Fura 2 fluorescence was measured
in cell populations with a Perkin-Elmer LS50B fluorescence
spectrophotometer. This instrument is equipped with a rotating filter wheel that can be used to alternate 340 and 380
nm excitation at approximately 50 Hz. A coverslip was
mounted vertically at a 30° angle to the light path in a
cuvette that was continuously perfused with media. A fourway valve mounted just above the cuvette allowed for rapid
switching of solutions; replacement of the medium bathing
the cells has a half-time of ⬃20 s. The excitation light illuminated an area of ⬃30 mm2 on the coverslip for recording of
fluorescence from several thousand cells. Background fluorescence was determined at the end of the experiment by
quenching the fura 2 fluorescence for 10–15 min in the
presence of 5 ␮M ionomycin and 6 mM MnCl2 in Ca2⫹-free
medium. After background fluorescence was subtracted, the
340- and 380-nm value ratios were calculated and calibrated
in terms of [Ca2⫹]i.
As described previously (4), calibration of fura 2 fluorescence in terms of [Ca2⫹]i utilized solutions of known Ca2⫹
concentration to construct a standard curve. The Ca2⫹ concentration of the standard solutions was calculated using
software (MaxChelator 6.60) that accounts for binding of
Ca2⫹ to each constituent of the solution.
For analysis of fluorescence ratios recorded from cells, the
equation [Ca2⫹]i ⫽ Kd ⫻ ␤ ⫻ [(R⫺Rmin)/(Rmax⫺R)] was used,
where Kd is the dissociation constant, ␤ is the ratio of fluorescence values for Ca2⫹-free to Ca2⫹-bound fura 2 measured
at the 380 nm excitation wavelength, R is the ratio of the
fluorescence intensities measured at 340 and 380 nm, and
Rmin and Rmax are the minimum and maximum values,
respectively, of the fluorescence ratio (R). The equation was
fit to the standard curve (using SigmaPlot software; SPSS)
and used to convert ratios (R) into [Ca2⫹]i values (10). In situ
calibration of fura 2 fluorescence by direct determination of
Rmin and Rmax from within cells yielded similar calibrated
values (not shown). Traces shown are representative of at
least three similar experiments.
Separation of [3H]arachidonic acid and [3H]phosphatidic
acid by thin layer chromatography. A7r5 cells were grown to
confluence in 60-mm petri dishes and labeled for 24 h with
[3H]arachidonic acid [AA; 1 ␮Ci in 3 ml of Dulbecco’s modified
Eagle’s medium (DMEM) supplemented with 3.5% fetal bovine serum at 37°C]. The cells were washed twice with
control medium supplemented with 0.5% fatty acid-free BSA
and preincubated for an additional 37.5 min in control medium supplemented with 0.1% fatty acid-free BSA in the
presence or absence of 5 ␮M ONO-RS-082. The medium was
removed and the cells were then treated with 100 pM AVP ⫾
5 ␮M ONO-RS-082 in 1.5 ml of medium for 27 min at 37°C.
At the end of this incubation, the medium was removed and
extracted according to the methods of Bligh and Dyer (1). The
extracted lipids from the medium were dried in a SpeedVac
rotary evaporator, resuspended in chloroform, and spotted on
Whatman 60A-LKD thin layer chromatography (TLC) plates.
The plates were developed in a solvent system of ethyl acetate, hexane, acetic acid, and water (85:35:15:90 parts, respectively). The locations of the lipids on the plate were
detected using phosphomolybdic acid, and bands corresponding to the locations of unlabeled AA and phosphatidic acid
(PA) were scraped into scintillation vials containing 1 ml of
methanol. After at least 30 min in methanol, 5 ml of xylenebased scintillant was added and the samples were counted in
an LKB-1209 liquid scintillation counter. Results of at least
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ROLE OF PLD IN AVP-STIMULATED CA2⫹ SPIKING IN A7r5 CELLS
AA-861, or 100 nM to 5 ␮M baicalein), and cytochrome
P-450 (5–20 ␮M methoxsalen or 1 mM 1-aminobenzotriazole) were ineffective in preventing AVP-stimulated Ca2⫹-spiking activity although some had modest
stimulatory effects (data not shown). Ketoconazole (10
␮M), a widely used cytochrome P-450 inhibitor,
slightly inhibited AVP-induced Ca2⫹ spiking, but this
concentration also inhibited [Ca2⫹]i changes induced
by increasing extracellular K⫹ concentration, which
suggests a nonspecific effect on Ca2⫹ channels. These
results indicate that metabolism of AA is unlikely to be
required for stimulation of Ca2⫹ spiking.
Lysophospholipids. Lysophospholipids are also products of PLA2 activity. In a series of experiments, several lysophospholipids were tested for effects on Ca2⫹
spiking. None of the compounds tested (lysophosphatidylethanolamine, lysophosphatidylcholine, lysophosphatidylserine, lysophosphatidylinositol, lysophosphatidic acid, and lysophosphatidylglycerol) induced Ca2⫹
spiking at concentrations between 1 and 100 ␮M (not
shown). In general, no effect on [Ca2⫹]i was observed
except at the highest concentrations tested (100 ␮M),
which in some cases (i.e., lysophosphatidylethanolamine and lysophosphatidylcholine) induced a steady
increase in [Ca2⫹]i that was reversed by washing away
the compound.
Phospholipase inhibition by ONO-RS-082. AVPstimulated AA release was previously assessed by labeling cells with [3H]AA and then measuring the radioactivity released into the medium after exposure to
Fig. 2. Inhibition of phosphatidic acid (PA) formation but not AA
release by ONO-RS-082. TLC was used to separate lipids released
into the medium in response to 100 pM AVP in A7r5 cells labeled for
24 h with [3H]AA. n, Number of experiments (in triplicate); *significantly different from control, P ⬍ 0.05.
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Fig. 1. Arachidonic acid (AA) does not stimulate Ca2⫹ spiking.
A: intracellular Ca2⫹ concentration ([Ca2⫹]i) recording from a fura
2-loaded A7r5 cell monolayer treated with varying concentrations
of AA (hatched boxes at top of panel indicate duration of exposure
to each of the indicated AA concentrations). B and C: cells treated
with 100 pM AVP (filled box, B) were washed for 10 min and then
exposed to 10 ␮M AA (filled box, C). Similar results were obtained
when AVP or AA were applied in reverse order (not shown).
AVP (2). Similar methods were used by Ito and colleagues (11) who also found that the release of 3H
radioactivity was stimulated by very low concentrations of AVP (EC50 ⬃ 50 pM). However, reports that
AVP stimulates PLD in VSM [including A7r5 cells (13,
15, 23)] led us to investigate whether some of the
radioactivity may be in the form of PA (the product of
PLD), which may contain the labeled fatty acyl moiety.
We used TLC to separate the lipid products and then
determined the radioactivity that comigrates with purified AA or PA standards. The results are shown in
Fig. 2 and indicate that ONO-RS-082 does not inhibit
AVP-stimulated AA release (open bars), but does inhibit PA formation (closed bars). These findings suggest that ONO-RS-082 inhibits AVP-stimulated PLD
rather than PLA2 activation.
The stimulation of PA formation by AVP might be
due to PLD activation or to phosphorylation of diacylglycerol. To unequivocally demonstrate activation of
PLD (6), A7r5 cells were labeled with [3H]palmitic acid
and then treated with AVP in the presence of 1-butanol. Under these conditions, only PLD activation will
lead to an increase in the formation of 3H-labeled
phosphatidylbutanol. As shown in Fig. 3, [3H]phosphatidylbutanol was significantly increased by 500 pM
AVP, and a larger increase occurred with maximal
AVP stimulation (100 nM AVP). Full concentrationresponse curves for AVP-stimulated phosphatidylbutanol formation (see Fig. 3, inset) revealed a half-maximal stimulation at an AVP concentration of 1.09 ⫾ 0.38
nM (n ⫽ 3).
Stimulation of Ca2⫹ spiking by PLD. Exogenous PA
added to the medium at concentrations from 10 nM to
ROLE OF PLD IN AVP-STIMULATED CA2⫹ SPIKING IN A7r5 CELLS
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10 ␮M did not stimulate Ca2⫹ spiking (not shown).
However, we also examined whether cleavage of endogenous phospholipids by PLD could stimulate Ca2⫹
spiking. We tested a bacterial PLD previously shown
by Jones and colleagues (13) to stimulate PA formation
to a similar level as AVP in A7r5 cells. As shown in Fig.
4, PLD (2.5 U/ml) produced a Ca2⫹-spiking response
that was indistinguishable from AVP (100 pM). Mean
spike amplitude was 233 ⫾ 3 nM for 100 pM AVP and
231 ⫾ 1.2 nM for 2.5 U/ml of PLD; spike frequencies
were 5.0 and 4.6 min⫺1, respectively. Bacterial PLA2
(2.5 U/ml) had no effect on Ca2⫹ spiking (not shown).
1-Butanol prevents AVP-stimulated Ca2⫹ spiking.
Butanol has been used by many groups to evaluate
PLD activity because it participates in a transphosphatidylation reaction that diverts PLD away from the
production of PA and toward the production of phosphatidylbutanol. 1-Butanol (0.2% vol/vol; ⬃22 mM)
Fig. 4. Stimulation of Ca2⫹ spiking by PLD. PLD type VII (2.5 U/ml)
added to the medium stimulated Ca2⫹ spiking (bottom). This effect is
indistinguishable from the effect of 100 pM AVP (top).
completely prevented AVP-stimulated Ca2⫹ spiking
(mean spike amplitude was 199 ⫾ 6 nM in AVP alone,
and frequency was 4.8 ⫾ 0.2 min⫺1 with no spiking
Fig. 5. 1-Butanol inhibits AVP- but not BaCl2-stimulated Ca2⫹ spiking. A7r5 cells were stimulated with AVP (100 pM; top) or BaCl2 (1
mM; bottom) in the absence (left) or presence (right) of 0.2% 1-butanol. Note that estimated [Ca2⫹]i values in bottom panels have not
been corrected for effects of Ba2⫹ on fura 2 fluorescence. Uncalibrated fluorescence ratios for spike peaks were 3.03 ⫾ 0.03 for 2 mM
BaCl2 alone and 3.02 ⫾ 0.01 for BaCl2 ⫹ 0.2% 1-butanol. Mean
frequency of spiking was 3.2 ⫾ 1.2 min⫺1 for BaCl2 alone and 4.3 ⫾
1.3 min⫺1 for BaCl2 ⫹ 1-butanol.
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Fig. 3. Phospholipase D (PLD) activation by AVP. PLD activity was determined by measuring [3H]phosphatidylbutanol ([3H]PBut) formation in response to AVP in the presence of 1-butanol (see MATERIALS AND METHODS).
Results from three experiments performed in duplicate are shown. *Significant difference from control; P ⬍ 0.001.
Inset: a representative concentration-response curve for AVP-stimulated [3H]PBut formation. [AVP], AVP concentration. Maximum stimulation based on a curve fit of the data to the equation y ⫽ a ⫻ x/(b ⫹ x) was set at 100%.
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ROLE OF PLD IN AVP-STIMULATED CA2⫹ SPIKING IN A7r5 CELLS
Fig. 7. PKC inhibition or downregulation does not prevent PLD
activation by AVP. [3H]phosphatidylbutanol formation was measured in A7r5 cells treated with 500 pM AVP in the presence of 0.2%
1-butanol. Where indicated, PMA pretreatment was for 24 h with 1
␮M PMA before AVP treatment; Ro-31-8220 was present for 1 h
before and during AVP treatment. Summarized results from seven
experiments performed in duplicate are shown. *Significant difference from controls treated with 1-butanol alone, P ⬍ 0.01.
activator of PLD or a downstream effector of PLD in a
variety of cell types (6, 7). In a previous study, we found
that AVP-stimulated Ca2⫹ spiking is prevented by the
PKC inhibitor Ro-31-8220 or by downregulation of
PKC isoforms by prolonged pretreatment with 4␤phorbol-12-myristate 13-acetate (PMA; see Ref. 8).
AVP-stimulated PLD activity was not inhibited by
PMA pretreatment or by the PKC inhibitor Ro-31-8220
(1 ␮M; see Fig. 7), which suggests that PKC activation
is downstream rather than upstream in the AVP signaling cascade.
DISCUSSION
Fig. 6. 2-Butanol does not inhibit AVP-stimulated Ca2⫹ spiking or
act as a substrate for transphosphatidylation. A: a population of
A7r5 cells was pretreated for 5 min with 0.2% 2-butanol then treated
with 100 pM AVP in the presence of 0.2% 2-butanol. B: a different
population of cells was treated with 100 pM AVP in the presence of
1-butanol (0.2%). C: [3H]phosphatidylbutanol formation was measured in A7r5 cells treated with or without AVP (500 pM) in the
presence of 0.2% 1-butanol or 0.2% 2-butanol. Results from two
experiments performed in duplicate are shown. *Significant difference from controls treated with 1-butanol alone, P ⬍ 0.01.
The results from the present study suggest that
AVP-stimulated PA formation by PLD is both necessary and sufficient to stimulate Ca2⫹ spiking in A7r5
cells. This finding may provide an important link between PLD and vasoconstrictor activity. PLD activity
has been demonstrated in many cell types including
VSM, where among the known activators of PLD are
the vasoconstrictor hormones AVP, ANG II, norepinephrine, and endothelin. The present findings expose
a novel signal-transduction pathway in which PLD
activation triggers Ca2⫹ signals that may underlie the
potent vasoconstrictor effects of AVP as well as other
vasoconstrictor hormones.
Although we previously suggested that PLA2 might
mediate AVP-stimulated Ca2⫹ spiking (2), further investigation has led us to the conclusion that PLA2 is
not the primary player in this signaling pathway. To
summarize: 1) AA release was only slightly increased
by 100 pM AVP and this effect was increased rather
than inhibited by ONO-RS-082, whereas AVP-stimulated PA formation was completely inhibited by ONORS-082; 2) AA and lysophospholipids (the products of
PLA2) were ineffective in stimulating Ca2⫹ spiking; 3)
agents that inhibited PLD-catalyzed PA formation
(ONO-RS-082 or 1-butanol) also prevented AVP-stim-
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observed in the presence of 0.2% of 1-butanol in four
experiments; see Fig. 5, top right). This concentration
of butanol did not apparently disrupt the cell membrane or adversely affect the Ca2⫹ homeostatic mechanisms because it did not affect resting [Ca2⫹]i levels or
the increase in baseline [Ca2⫹]i in response to AVP (see
Fig. 5, top right) or prevent stimulation of Ca2⫹ spiking
by BaCl2 (see Fig. 5, bottom right). BaCl2 likely stimulates Ca2⫹ spiking by inhibiting K⫹ channels, a mechanism that may be an element of AVP signaling downstream of PLD activation (21). 2-Butanol, a butanol
isomer that does not participate in the transphosphatidylation reaction (5), did not prevent AVP-stimulated
Ca2⫹ spiking (spike frequency averaged 4.8 min⫺1 in
100 pM AVP, and 6.1 min⫺1 in AVP ⫹ 2-butanol; see
Fig. 6, A and B). Phosphatidylbutanol formation was
not stimulated by AVP in the presence of 2-butanol (see
Fig. 6C).
Protein kinase C and PLD activation. Protein kinase
C (PKC) has been implicated as either an upstream
ROLE OF PLD IN AVP-STIMULATED CA2⫹ SPIKING IN A7r5 CELLS
tors (16, 22) and provides a potential link between AVP
binding and PLD activation by small molecular weight
G proteins.
Despite the paucity of information on the mechanisms of activation of PLD in vascular smooth muscle
by G protein-coupled agonists, it is clear that a variety
of vasoconstrictors activate PLD. In some instances,
PLD activation by AVP has been demonstrated to occur
with greater potency than the PLC-mediated formation of Ins(1,4,5)P3 (17), which suggests a qualitative
as well as quantitative concentration-dependent pattern of second-messenger formation. If different signaling pathways are activated over different ranges of
agonist concentration, the pathways may produce different physiological endpoints. Very high concentrations of vasoconstrictor hormones are known to lead to
vascular remodeling involving hypertrophy and/or hyperplasia of smooth muscle cells. These vascular alterations have been implicated in pathological processes
such as the development of atherosclerosis and hypertension. From a teleological point of view, it would be
appropriate to provide for systemic regulation of
smooth muscle contractility without stimulating
events that may lead to vascular remodeling. Our
present findings lead us to speculate that systemic
concentrations of AVP in the picomolar range (below
the concentrations required to induce vascular remodeling) may modulate the frequency of Ca2⫹ spiking in
VSM of resistance vessels by activation of PLD and
thereby regulate tissue perfusion and peripheral resistance.
The authors thank Matt Hammoudeh and John Barakat for technical assistance.
This work was supported by the Eugene J. and Elsie E. Weyler
Endowment for Clinical Cardiology Research, the John and Marion
Falk Trust for Medical Research, and the National Heart, Lung, and
Blood Institute (Grant 1R01 HL-60164-01A1).
Present address for A. J. Shiels: Department of Internal Medicine,
Washington University School of Medicine, Barnes-Jewish Medicine
Clinic South Campus, St. Louis, MO 63110.
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ulated Ca2⫹ spiking; and 4) exogenous PLD, which has
been shown previously to stimulate PA formation (13),
stimulated Ca2⫹ spiking, whereas exogenous PLA2 did
not.
The postulated role of PLD in stimulation of Ca2⫹
spiking implies a temporal sequence in which PLD
activation would precede downstream signaling steps
and the initiation of Ca2⫹ spiking. Unfortunately, the
methods employed to detect PLD activation are not
sensitive enough to detect the relatively modest increase in PLD activity at low AVP concentration at
early time points. However, we do not feel that our
inability to detect these rapid changes in PLD activation necessarily diminishes the potential importance of
PLD in the cellular responses observed. It is increasingly apparent that signaling cascades operate within
subcellular microdomains where minute changes in
the biochemical environment can lead to profound cellular responses. Sophisticated methods have recently
been developed to measure tiny subcellular [Ca2⫹]i
changes (e.g., Ca2⫹ sparks) that are believed to play
important roles in smooth muscle physiology. These
Ca2⫹ sparks were impossible to resolve previously by
methods that detected the larger global [Ca2⫹]i
changes induced by larger stimuli. Similarly, more
sophisticated techniques for measuring PLD activity in
real time may be required to detect the earliest subcellular activity stimulated by physiological concentrations of AVP.
It remains to be determined how binding of AVP to
vasopressin V1a receptors activates PLD. Heterotrimeric G proteins have been implicated in PLD activation. For example, ␤␥-subunits and G␣12 have recently
been postulated to be involved in ANG II-stimulated
PLD activity in VSM (25), and G␣13 (19) has been
implicated in activation of PLD in nonmuscle cells. At
least two isoforms of PLD (PLD1 and PLD2) are expressed in mammalian cells. Both of these isoforms are
expressed in A7r5 cells (Ref. 9 and K. L. Byron, unpublished observation). Whereas evidence from several
laboratories has suggested that PLD1 may be regulated by numerous factors including small molecular
weight G proteins (e.g., ADP ribosylation factor and
RhoA), PKC, and tyrosine kinases, regulation of PLD2
is still poorly characterized (for reviews, see Refs. 6
and 7).
Interestingly, the pressor responses of spontaneously hypertensive rats to AVP were reportedly inhibited by the cholesterol-lowering drug lovastatin (12),
and this effect was associated with a decreased expression of small molecular weight G proteins (Ras and
Rho) but not heterotrimeric G proteins (Gs, Gi, or Gq).
Activation of PLD by G protein-coupled receptors has
been recently reported to involve a specific domain
within the heptahelical receptor (14). The Asn-Pro-XTyr motif in the seventh transmembrane domain of
rhodopsin family receptors that couple to PLD activation is postulated to form a functional complex with the
small molecular weight G proteins, ARF and RhoA.
This motif is also present in the seventh transmembrane domain of human and rat V1a vasopressin recep-
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